obap, a new family of oil body-associated proteins author to whom

1

Running head:

OBAP, a New family of Oil Body-Associated Proteins

Author to whom all correspondence should be sent:

Carlos M Vicient

Department of Molecular Genetics

Centre for Research in Agricultural Genomics CRAG (CSIC-IRTA-UAB-UB)

Campus UAB, Bellaterra (Cerdanyola del Vallès) 08193 – Barcelona (Spain)

Telephone +34 935636600

Email: [email protected]

Research Area most appropriate for the paper:

Cell Biology

Plant Physiology Preview. Published on January 9, 2014, as DOI:10.1104/pp.113.233221

Copyright 2014 by the American Society of Plant Biologists

www.plantphysiol.orgon January 29, 2018 - Published by Downloaded from Copyright © 2014 American Society of Plant Biologists. All rights reserved.

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2

Title of article:

The evolutionary conserved oil body associated protein OBAP1 participates in

the regulation of oil body size

All authors' full names

Ignacio López-Ribera1, José Luis La Paz1, Carlos Repiso1, Nora García1, Mercè

Miquel1, María Luisa Hernández2, José Manuel Martínez-Rivas2, Carlos M

Vicient1

Institution address

1 Department of Molecular Genetics. Centre for Research in Agricultural

Genomics CRAG (CSIC-IRTA-UAB-UB). Campus UAB, Bellaterra (Cerdanyola

del Vallès) 08193 – Barcelona (Spain)

2 Department of Biochemistry and Molecular Biology of Plant Products. Instituto

de la Grasa (CSIC), 41012-Sevilla (Spain)

A one-sentence summary:

A new family of oil body associated proteins present in plants, fungi and

bacteria are involved in determining oil body size and shape.



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Financial source:

This work was supported by the Spanish Ministry of Science and Innovation

(BIO2007-64791 and 2007-20I037), the CONSOLIDER-INGENIO programme

(CSD2007-00036), and the Xarxa de Referencia en Biotecnologia of the

Generalitat de Catalunya. M.L.H. was the recipient of JAE-Doc contract from

CSIC.

corresponding author; e-mail address

Carlos M Vicient, [email protected]



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ABSTRACT

A transcriptomic approach has been used to identify genes predominantly

expressed in maize (Zea mays L.) scutellum during maturation. One of the

identified genes is obap1, which is transcribed during seed maturation,

predominantly in the scutellum, and its expression decreases rapidly after

germination. Proteins similar to OBAP1 are present in all plants, including

primitive plants and mosses, and in some fungi and bacteria. In plants, obap

genes are divided in two subfamilies. Arabidopsis (Arabidopsis thaliana L.)

genome contains five genes coding for OBAP proteins. Arabidopsis OBAP1a

protein is accumulated during seed maturation and disappears after

germination. Agroinfiltration of tobacco epidermal leaf cells with fusions of

OBAP1 to YFP and immunogold labeling of embryo transmission electron

microscopy sections showed that OBAP1 protein is mainly localized in the

surface of the oil bodies. OBAP1 protein was detected in the oil body cellular

fraction of Arabidopsis embryos. Deletion analyses demonstrate that the most

hydrophilic part of the protein is the responsible of the oil body localization,

which suggests an indirect interaction of OBAP1 with other proteins in the oil

body surface. An Arabidopsis mutant with a T-DNA inserted in the second exon

of the obap1a gene and an RNA interference line against the same gene

showed a decrease in the germination rate, a decrease in seed oil content,

changes in fatty acid composition and their embryos have few, big and irregular

oil bodies, compared to wild type. Taken together, our findings suggest that

OBAP1 protein is involved in the stability of oil bodies.



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INTRODUCTION

The scutellum is a shield-like structure surrounding the embryo axis that occurs

in all grass species, including early-diverging taxa, and is generally considered

as functionally equivalent to the cotyledon (Negby, 1984; Vernoud et al., 2005).

The scutellum plays an important role in the hydrolysis and transport of

endosperm stored substrates during germination, but the scutellum itself also

accumulates part of the seed reserves, in special lipids (Tzen and Huang, 1992;

Subbarao et al., 1998; White and Weber, 2003). Maize (Zea mays L.) scutellum

only represents 11% of the mass of the kernel, but accumulates 90% of the oil,

20% of the storage proteins, 10% of the sugars and 90% of the phytate

(Watson, 2003) and is the main source of vitamins and minerals of the kernel

(Mazzolini et al., 1985; Lombi et al., 2011).

Triacylglycerols (TAGs) in seeds accumulate in special cytoplasmatic organelles

called oil bodies (OBs) (Murphy, 2001) which consists in a hydrophobic central

core of neutral lipids, such as TAGs, surrounded by a monolayer of amphipathic

phospholipids, glycolipids and/or sterols, with a series of proteins bound to the

surface of the OB (Purkrtova et al., 2008). OBs are also present in Animals,

Fungi and Prokaryotes (Yang et al., 2012). Although the main role of OBs is to

accumulate nutrients, increasing data indicate that in Eukaryotic cells OBs are

also involved in other roles as lipid and protein trafficking between organelles

(Raposo and Stenmark, 2008). Proteomic analyses suggest that OBs can serve

as transient storage depots for proteins that lack appropriate binding partners in

the cell and may provide a general cellular strategy for handling excess proteins

(Cermelli et al., 2006). OBs also seem to be involved in the protection of plant

embryos against freeze (Shimada et al., 2008).

A limited number of proteins are associated with OBs. Each type of organisms

contains a specific set of them, for example, perilipins in mammal cytosolic OBs

(Kimmel et al., 2010), 19 proteins in fungi (Grillitsch et al., 2011), and phasins in

bacterial OBs (Yang et al., 2012). The most abundant plant proteins associated

with OBs are oleosins (Capuano et al., 2007; Huang et al., 2009). Oleosins are

tightly associated with OBs due to the presence of a highly hydrophobic central



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“core” composed of 70 to 80 hydrophobic and non-polar amino acids (Murphy,

2001). In the center of this hydrophobic stretch lies a conserved motif containing

three Pro residues that are crucial for the OB localization of oleosins (Abell et

al., 1997). Oleosins maintain the integrity of the OBs and regulate their size by

preventing them from coalescence (Tzen and Huang, 1992). Caleosins are a

second group of plant proteins associated to OBs which also contain a

hydrophobic core of about 30 amino acids and a E-F hand calcium-binding motif

in its C-terminal end (Chen et al., 1999; Frandsen et al., 2001). Caleosins seem

to have a role in OB degradation during germination (Poxleitner et al., 2006).

Another plant OB associated proteins are steroleosins, characterized by the

presence of a NADPH-binding region and a soluble sterol-binding

dehydrogenase domain (Lin et al., 2002). Oleosins are not present in green

algae, but a lipid droplet surface protein (LDSP) was identified in the algae

Nannochloropsis sp with structural similarities to other OB associated proteins

(Vieler et al., 2012) and a major lipid droplet protein was also identified in

Chlamydomonas and other green algae species (Moellering and Benning,

2010; Peled et al., 2011). Proteomic analysis in different plants and algae

identified a wide variety of proteins associated to oil bodies (Jolivet et al., 2009;

Katavic et al., 2006; Nguyen et al., 2011; Tnani et al., 2011; Tnani et al., 2012).

Different roles have been proposed to the oil body associated proteins,

including OB biogenesis, stability, trafficking and mobilization.

In this study, we identified the genes predominantly expressed in the scutellum

during seed maturation compared to other parts of the seed. Among them, we

identified a gene coding for a protein of unknown function we called OBAP1.

Homologous genes are present in all plant species, and in some fungi and

bacteria. Here, we demonstrate that OBAP1 is associated to the OBs. Our

results demonstrate that OBAP1 is necessary to maintain the structure of the

OBs and for seed germination in Arabidopsis (Arabidopsis thaliana L.).



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RESULTS

Identification of maize genes predominantly expressed in scutellum

Scutellum samples were dissected from immature embryos 30 days after

pollination (dap) which corresponds to the embryo maturation stage marked by

the growth of the embryo and the accumulation of reserve substances (Vernoud

et al., 2005). RNA was extracted and a cDNA library constructed. EST clones

from the unamplified library were randomly picked and those with inserts larger

than 130 bp were sequenced from their 5’ends. Tables S1 and S2 list the 1,553

sequenced ESTs and their corresponding 789 genes, and include accession

numbers, functional category and best BLASTP match. Fifty five ESTs

corresponded to transposable elements (3,5%). 67% of the ESTs corresponding

to genes were singletons and the rest assembled into contigs of two to 37

sequences. Among the 18 largest contigs, composed of more than seven ESTs,

the majority corresponded to zeins. Considering all genes with an identified

function, 30% are involved in metabolism (25 in lipid metabolism) and 30% in

protein synthesis and processing (Table S3). In order to identify genes

specifically or predominantly expressed in the immature scutellum, 618 clones

corresponding to different genes were selected from the scutellum library (Table

S4). The inserts of the cDNA clones were PCR amplified and spotted onto nylon

membranes. Macroarray hybridization experiments were carried out in triplicate

with radio-labeled retrotranscribed probes synthesized with RNA extracted from

dissected embryo axis, endosperm and scutellum (30 dap). A gene was

designated as being predominantly expressed in scutellum when, after

normalization, the average signal intensity with scutellum probes was greater

than twice and a half that of embryo axis and of endosperm samples. Four

genes showed a significantly higher expression in scutellum (Table S4): two

genes coding oleosins (oleosin I and II), a gene involved in defense against

pathogens (antimicrobial peptide MBP-1) and a gene of undetermined function

(GRMZM2G044627). This last gene is represented by three clones in the

scutellum library, which indicates a moderately high level of expression. We

named it obap1 because, as we will show later, it encodes an Oil Body

Associated Protein.



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OBAP1 belongs to an evolutionary conserved family of proteins

The obap1 gene is located in chromosome 4 (155907185-155909704) and

contains one intron. Transcript sequence databases describe only one transcript

variant and the in silico expression database eFP Browser suggests that higher

expression occurs in mature embryo (Sekhon et al., 2011). The maize genome

contains two additional genes encoding proteins similar to OBAP1: obap2a

(GRMZM2G043521), located in chromosome 3 and 33% similar, and obap2b

(GRMZM2G107570), located in chromosome 8 and 36% similar. OBAP2A and

OBAP2B are 63% similar between them. The genes obap2a and obap2b also

contain one intron and the in silico data indicate that their highest expression

also occurs in mature embryo. A total of 235 protein sequences similar to maize

OBAP1 were identified in databases corresponding to 153 species, 165 of them

full-length sequences (Table S5). They corresponded to several plant species

including monocots, dicots, conifers, primitive plants, mosses and algae

(Liliopsida, Eudicotyledons, Conipherophyta, Filicophyta, Licopodiophyta,

Bryophyta and Chlorophyta), but also to some Fungi (Ascomycota,

Basidiomycota, Oomycetes and Zygomycetes) and some Prokaryotes

(Proteobacteria, Planctomycetes, Bacteroidetes and Actinonbacteria). Only one

sequence corresponding to an animal species was found (Acc. Num.

FG621146). This sequence corresponds to an EST collection obtained from

desiccated samples of the nematode Plectus murrayi (Adhikari et al., 2009).

However, it is unlikely that this sequence really corresponds to a Plectus

murrayi transcribed gene because similar genes are not present in the genomes

of any other nematode species, including the fully sequenced genome of

Caenorhabditis elegans, and because this sequence is 100% similar to a

peanut transcribed mRNA (Acc. Num. EE126736), which strongly suggest that it

corresponds to a contamination. In conclusion, according to the current

available information, obap genes are not present in animals. Whereas in plants

obap genes are present as small gene families, Fungi and Prokaryote genomes

contain only one copy, although not in all species. For example, Escherichia coli

and Saccharomyces cereviseae genomes do not contain obap genes. A

phylogenetic tree based on all the available OBAP full-length sequences shows



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that there are two OBAP families in plants, one in Fungi and one in Prokaryotes

(Figure 1; Figure S1). Both plant subfamilies contain genes from multiple

species and representatives from monocotyledons, dicotyledons, conifers and

primitive plants. In general, monocot plants contain three copies of the gene,

one from family 1 and two from family 2, and dicots, conifers and primitive

plants contain a variable number of genes distributed in the two subfamilies. For

example, Arabidopsis thaliana genome contains two genes of subfamily 1

(obap1a and obap1b) and three genes of subfamily 2 (obap2a, obap2b and

obap2c). Inside each group the sequences are distributed accordingly to the

species phylogeny (Figure S2).

Expression pattern of maize obap1 gene

qRT-PCR experiments revealed that at 30 days after pollination (dap) maize

obap1 mRNA is about 20 fold more abundant in the scutellum than in the

embryo axis, and about 100 times more in the scutellum than in the endosperm

(Figure 2A), confirming the macroarray results. No expression was detected in

leaves or roots. A similar pattern was obtained for oleosin1 gene (Figure 2B).

Expression was also examined in embryos at different stages of development.

In both genes the highest expression was found in maturing embryos, but

whereas obap1 have a maximum at about 35 dap (Figures 2C), oleosin1 have a

maximum about ten days earlier (Figure 2D). After imbibition, the mRNA

accumulation in embryos in both cases decreases rapidly (Figures 2E and F).

The obap1a gen in Brassicaceae

Five obap genes are present in the Arabidopsis thaliana genome. At1g05510

(AtObap1a) is the more similar to the maize obap1. According to Arabidopsis

microarray databases (Arabidopsis eFP Browser), AtObap1a is highly

expressed during seed maturation, its expression drops quickly after imbibition

and no expression was observed in other organs (Figure S3). An antibody was

produced against Arabidopsis OBAP1a and used in western blot analyses in

order to study protein accumulation in different organs and during seed

development and germination (Figure 3). AtOBAP1 protein is not present in



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seeds at early stages of development, but accumulated rapidly during seed

maturation (Figure 3A). Mature leaves and stems do not contain OBAP1a.

Protein accumulation decreases rapidly after imbibition, becoming undetectable

only 5 days after imbibition (Figure 3B).

Subcellular localization of OBAP1 protein

OBAP1 protein sequences do not contain any recognizable protein motif or

transmembrane helix that could give us clues about its function. Maize OBAP1

(LOC 100193701) and Arabidopsis OBAP1a (At1g05510) proteins Appeared in

proteomic analysis of maize embryos and Arabidopsis seeds and in both cases

their concentration increased during artifitial ageing (Rajjou et al., 2008; Xin et

al., 2011). Arabidopsis OBAP1a also appeared in a proteomic analysis focused

in the identification of proteins whose Tyr phosphorylation varied significantly in

seeds imbibed for 2 days in ABA (Ghelis et al., 2008). These proteomic analysis

did not provide any experimental data on the possible subcellular distribution of

AtOBAP1a, however, the protein is defined by Ghelis et al., (2008) as a

“putative lipoprotein”. Some of the homologous genes from Prokaryote species

are also described similarly in GeneBank, although no experimental data have

been published for none of them. In order to clarify OBAP1 subcellular

localization, the full-length ZmObap1 coding region was fused with the YFP

coding region in C-terminal under the control of 35S promoter and analyzed by

confocal laser scanning microscopy after transient expression by agroinfiltration

in Nicotiana benthamiana leaf epidermal cells. Confocal microscopy revealed

YFP fluorescence in small organelles distributed in the cytoplasm (Figure 4A).

Fluorescence obtained with YFP alone shows a more uniform distribution in the

cell (Figure 4B). This punctuate distribution was similar to the distribution of

OBs in tobacco cells as can be seen after Nile Red treatment that stains neutral

lipids and it has been used to localize oil bodies (Huang et al., 2009). Used in

tobacco leaf epidermal cells, Nile Red stains spherical lipid-based particles

distributed in the cytoplasm (Figure 4C). When tobacco leaf epidermal cells

agroinfiltrated with OBAP-YFP construct (Figure 4D) were stained with Nile Red

(Figure 4E), both signals mainly co-localized (Figure 4F), indicating that the

punctuate structures in which OBAP1-YFP localizes correspond to OBs. Finally,



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we carried out co-expression experiments to verify the co-localization of OBAP1

with the well-known OB associated protein OLEOSIN2 (Figures 4G-I). As

expected, OLEOSIN2-CFP was localized in spherical bodies (Figure 4G) and

part of the OBAP-YFP protein co-localized with OLEOSIN2 (Figure 4H).

However, the ectopic expression of OLEOSIN2-CFP induces that part of the

OBAP1-YFP protein appeared to be located in aligned small spots (Figure 4I)

that do not contain OLEOSIN2-CFP (Figure 4I).

We used the anti-OBAP1 antibody in order to obtain further evidence of the

association of OBAP1a to OBs. First, immunogold transmission electron

microscopy was used to determine AtOBAP1a subcellular localization in

rapeseed mature embryos. Gold-labeling was observed predominantly in the

surface of the OBs (Figure 5). Second, we used the flotation centrifugation

method (Katavic et al., 2006) in order to isolate the OB-enriched protein fraction

from rapeseed dry seeds. The OB layer, the total lysate and the cytosolic

fraction were delipidated, precipitated, separated by SDS-PAGE and blotted

onto a nitrocellulose membrane for inmunodetection (Figure 6). Although the

antibody raised against Arabidopsis OBAP1a recognized proteins present in the

OB protein fraction, demonstrating the association of OBAP with the OBs, it

also recognized proteins present in the cytosolic fraction.

Confocal and transmission electron microscopy experiments indicated that

OBAP1a is mainly located in the OBs. However, subcellular fractioning

experiment raised some doubts respect this subcellular distribution. The

association to OBs of other plant proteins like oleosins and caleosins are known

to be due to the presence of a long central highly hydrophobic domain. Oleosins

have a central hydrophobic domain of about 70 residues (Figure S4A;

Sarmiento et al., 1997), caleosins of about 30 residues (Figure S4B; Naested et

al., 2000) and steroleosins of about 40 residues (Figure S4C; Lin et al., 2002).

These proteins also contain a characteristic proline-rich motif. OBAP1 proteins

do not contain a proline-rich motif and, although hydropaty plots show some

hydrophobic regions, they are short (less than 20 residues) and not as much

hydrophobic as in the other three plant oil body associated proteins (Figure 7A).

In order to determine if these short hydrophobic regions are the responsible of



12

the OB localization of OBAP1 we divided the obap1 coding region in two parts

and fused them to the yfp coding region: The N-terminal part (N-OB) contains

the hydrophobic regions and the C-terminal part (C-OB) contains the most

hydrophilic part of the protein (Figure 7B). When introduced into Nicotiana

benthamiana leaf epidermal cells, the N-terminal part shows a sub-cellular

distribution similar to YFP alone (Figure 7C-D) whereas the C-terminal part

shows a punctuate distribution characteristic of the OB distribution (Figure 7E).

These results suggest that OBAP1 does not interact with the central lipidic

domain of the OBs and that OB localization is due to another type of interaction

as, for example, the interaction with another OB associated protein.

Insertion mutants and RNAi lines for AtObap1a gene

We used insertional mutagenesis and RNA interference (RNAi) approaches to

determine the biological function of Arabidopsis OBAP1a. Public collections of

T-DNA insertion in Arabidopsis were screened and we have found two lines

containing T-DNA insertions in AtObap1a gene. Seeds of these homozygous

mutant lines were obtained from NASC T-DNA collection. One insertion was

located in the second exon (Figure 8A) and we designated it AtObap1a-1 (1A1).

The second insertion was located about 500 bp upstream the ATG and we

called it AtObap1a-2 (1A2). We have also used the RNA interference (RNAi)

method to reduce AtOBAP1a expression. We have produced an RNAi line

specific of AtObap1a gene we called Ag1. Immunoblot analyses using anti-

AtOBAP1a antibody showed that, whereas the abundance of OBAP1a protein

in 1A2 seeds is not altered, the seeds of the 1A1 line do not contain detectable

amounts of OBAP1a protein and the seeds from Ag1 line contain a reduced

quantity (Figure 8B). Whereas 1A2 seeds germinated similarly to wild type, Ag1

seeds germinated poorly (about 50% respect wild type) and mutant 1A1 seeds

germinated less than 2% respect wild type (Figure 8C). After germination, no

difference in seedling development were observed between Ag1 or 1A1 lines

and wild type. No significant differences were observed in the weight of the

seeds of the different lines.



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To investigate whether alterations in AtOBAP1a protein deposition would affect

the synthesis of storage lipids, we performed biochemical analyses of seed TAG

content and FA composition. When compared to wild-type we observed no

differences in the total amount of TAG in 1A2 and Ag1 seeds, but a significant

reduction in 1A1 seeds, which contain about 60% TAG respect to wild-type

(Figure 8D). Although there are quantitative differences in the accumulation of

TAG, qualitative analyses of FA composition showed only minor statistically

significant differences when compared to wild type (p<0.05 by Student’s t-test;

Figure 8E). Whereas 1A2 seeds did not show any significant differences in

overall FA profiles in TAG respect to wild type, the seeds of the 1A1 and Ag1

lines contain significantly more linoleic acid (18:2) (20% increase in 1A1 and

12% increase in Ag1) and less eicosenoic acid (20:1) (10% reduction in 1A1

and 4% reduction in Ag1).

We conducted microscopy analyses in order to investigate the possible effects

of different AtOBAP1a accumulation on the morphology of OBs. First, we

extracted embryos from mature seeds of each line, stained them with Nile Red

(which specifically stains neutral lipids) and observed them in the confocal

microscope (Figure 9A-C). Whereas wild-type embryos showed a uniform

distribution of relatively small OBs (Figure 9A), embryos from 1A1 (Figure 9B)

and Ag1 (Figure 9C) contained bigger OBs with an irregular shape and located

in the central part of the cell. In order to obtain a more precise image of these

changes in OB structure the embryos were also observed using electron

microscopy (Figure 9D-I). Compared to wild type (Figure 9D and E), the oil

bodies in embryos of 1A1 mutant (Figure 9F and G) and Ag1 RNAi line are

enlarged and irregularly shaped (Figure 9H and I). The effect is more intense in

the 1A1 insertion line compared to Ag1.

DISCUSSION

Maize kernels contain about 4,5% oil and about 90% is accumulated in the OBs

of the scutellar parenchymal cells (Watson, 2003). In consequence, the maize

scutellum could be considered as a good source in order to identify new genes

and proteins involved in the synthesis and accumulation of storage lipids in



14

plants. In kernels of 30 days after pollination (dap) the scutellar cells did not

further divide and they only increase in size and accumulate storage molecules,

including oil (Bommert and Werr, 2001). The peak of oleosin gene expression

occurs about 25 dap (Figure 2D). Functional categories of the expressed genes

in 30 dap scutellum showed a predominance of metabolic functions and a low

representation of genes involved in cell division and development. Genes

coding for storage proteins and for proteins involved in protein synthesis and

processing are also highly represented, reflecting the fact that scutellum also

accumulates storage proteins. A transcriptomic analysis of dissected barley

embryos at a similar developmental stage also showed that ribosomal proteins

and other components of the translation machinery are highly expressed in the

scutellum (Potokina et al., 2002). We identified four genes predominantly

expressed in scutellum and two of them encode oleosins, which is coherent with

the oil storage function of this organ. Another gene encodes an antimicrobial

peptide MBP-1. Pathogens could try entering into the embryo through the

endosperm, and scutellum seems to play a protective role against them.

Accordingly, some genes involved in pathogen defense have been described to

be expressed in scutellum: a barley gene coding for a beta-1,-3-glucanase, a

maize gene coding for a ribosome-inactivating protein (RIP) and the maize and

wheat cystatin genes (Jutidamrongphan et al., 1991; Guo et al., 1999, Corre-

Menguy et al., 2002).

No clues were available for the role of the protein encoded by the fourth

identified gene. We have demonstrated here that the protein is associated to oil

bodies and we called it OBAP1 (Oil Body Associated Protein 1). All the available

expression data (generated by us or available in expression databases) indicate

that obap genes are predominantly expressed during embryo development and

their expression pattern is very similar to the oleosin genes. Fusions of OBAP1

to fluorescent protein tags demonstrate that OBAP1 co-localizes with OBs in N.

benthamiana leaf epidermal cells, which contains bona fide OBs (Wahlroos et

al., 2003). These results were confirmed by immunolocalization experiments of

OBAP1a in electronmicroscopy samples of rapeseed mature embryos. Although

oleosins are the most abundant plant OB associated proteins, they are not the

unique. Caleosins (Chen et al., 1999) and steroleosins (Lin et al., 2002) are also



15

localized in OBs, and the proteomic analyses on maize scutellum and Brassica

seeds suggest that other less abundant OB associated proteins must exist

(Katavic et al., 2006; Hajduch et al., 2007; Tnani et al. 2012).

An apparently contradictory result was obtained in subcellular fractioning

experiment in which OBAP1 proteins appears to be located in both the OB and

the soluble fractions. However, something similar happens with oleosins, the

more characteristic plant OB proteins, because about 5-10% of rapeseed

oleosins were invariably targeted to the non-OB fraction in similar experiments

(Sarmiento et al., 1997) and, similarly, a proportion of Arabidopsis caleosin can

be detected in the soluble fraction (Naested et al. 2000). Despite these, in situ

experiments demonstrate that both proteins, as well as OBAP1 protein, are

predominantly located in the surface of the OBs. The proportion of protein

located in the OB fraction is much lower in the case of OBAP1 compared to

oleosin and caleosin and this may reflect that OBAP1 attachment to OBs is

weaker. Oleosin and caleosin association to OBs is mediated by the presence

of a long central hydrophobic region, however, OBAP1 does not have long

hydrophobic regions and the fraction of the protein responsible of the interaction

is the C-terminal, corresponding to the most hydrophilic part of the protein.

These data suggest that probably the interaction of OBAP1 with OBs is not

mediated by a direct interaction of the protein with the central TAG core of the

OB. Alternatively, OBAP1 may interact with OBs in a similar manner as has

been suggested to MLDPs in Algae, that is, by the direct contact of its polar

residues with the phospholipids or with other components of the surface of the

OBs, like other proteins (Huang et al., 2013). Oleosins and caleosins contain a

signal peptide and are directed to the ER and trafficked to the surface of OBs

(Beaudoin and Napier 2002; Froissard et al. 2009). Unlike oleosins, OBAP1 do

not have a signal peptide, which suggests that initially it accumulates in the

cytoplasm rather than in ER, and this is also coherent with the hypothesis of a

superficial interaction of OBAP to the OBs once they have been formed.

Coherently to this hypothesis, the pick of expression of OBAP1 in maize

embryos occurs later than the pick of expression of oleosin.



16

When obap1 is co-expressed with oleosin2, part of the OBAP1 protein localizes

in linear structures resembling ER. Interestingly, overexpression of oleosin has

a similar effect on caleosin localization (DeDomenico et al., 2011). These results

suggest the possibility that OBAP1 interact with caleosins and not with oleosins.

Homology searches indicate that obap genes are present in all plant species,

including primitive plants, and in some Fungi and Prokaryotes. Oleosins are

only present in plants (Huang et al., 2009), but caleosins are present in plants

and some fungi (Murphy, 2005). Caleosin and OBAP are both present in, for

example, Aspergillus niger, Ustilago maydis, Magnaporthe grisea, Neurospora

crassa or Chaetomium globosum, but not in the yeast Saccharomyces

cereviseae. Further analyses will be necessary to determine this possible

interaction.

Irregularly expanded oil-containing structures appear when OBAP1 is not

present and seed germination rate and oil content decreases, whereas fatty

acid composition is altered. Oleosin mutants produce similar changes (Siloto et

al., 2006; Shimada et al., 2008) and authors suggested that OB expansion push

the nuclei to the cell periphery and may prevent them from playing their roles,

inducing cell degeneration and death (Shimada et al., 2008). On the other hand,

not only total oil content but also the average size of the OBs determines the

germination rate since large OBs reduce the accessibility of lipases to the TAGs

during germination (Siloto et al., 2006). From a qualitative point of view, the

suppression of OBAP1 has a significant effect on fatty acid preferences in TAG,

increasing 18:2 (linoleic acid) at the expense of 20:1 (eicosanoic acid). Similar

alterations in the oil content and fatty acid composition of TAGs were observed

in an Arabidopsis dgat1 (diacylglycerol acyltransferase 1) mutant in which TAG

biosynthesis is reduced (Katavic et al., 1995). All these data suggest that,

similarly to oleosins, OBAP1 protein is necessary to maintain OB integrity and

may also exert some influence on qualitative and quantitative aspects of TAG

accumulation in seeds.

Different studies have revealed that OBs, previously considered static storage

depots of cellular neutral lipids, may be more active organelles (Bartz et al.,

2007; Beller et al., 2006; Sato et al., 2006; Wan et al., 2007). Animal lipid



17

droplets interact with various cellular organelles (peroxisomes, glyoxisomes,

mitochondria, etc.) (Liu et al., 2007) and lipid droplets have also been

considered a refugee of proteins destined for destruction like toxins or viral

proteins, or proteins that will be used when conditions change like signaling

proteins (Welte, 2007). An interesting possibility is that OBAP mediates some of

these interactions. Supporting this idea, the Arabidopsis Interactome Mapping

Project, based on experimental yeast two-hybrid data, found that Arabidopsis

OBAP1a, OBAP1b and OBAP2c interact with TCP14 transcription factor.

Arabidopsis OBAP1b is also predicted to interact with other transcription factors

of the TCP family (TCP11 and TCP15) and with MARD1 (MEDIATOR OF ABA-

REGULATED DORMANCY 1)(Arabidopsis Interactome Mapping Consortium,

2011). If confirmed, these interactions open several new possible functions for

OBs and OBAP proteins in plants.



18

MATERIALS AND METHODS

Plant Materials and Plant Growth Conditions

Maize (Zea mays L cv W64A) were grown under standard conditions (16/8 hr at

26°C; 150 μmol m–2 s–1). Nicotiana benthamiana were grown in soil under long-

day conditions (16/8 hr at 22°C; 150 μmol m–2 s–1). Seeds from Brassica

carinata (N2-7399) and Glycine max (N85-2124) were obtained from USDA.

The following SALK Arabidopsis (Arabidopsis thaliana L.) insertion mutants

were acquired from the ABRC (http://abrc.osu.edu/): 1A1 (SALK_011689) and

1A2 (SALK_017397). The RNA interference line Ag1 was requested to

AGRIKOLA project (Systematic RNAi knockouts in Arabidopsis; www.

agrikola.org; Hilson et al., 2003).

For Arabidopsis germination assays, seeds were surface-sterilized for 2 min

with 75% ethanol, followed by 5 min in 1% NaClO solution, washed five times in

sterile distilled water, and then placed onto 0.8% w/v agar in Murashige-Skoog

(MS) medium pH 5.7. Plates were kept for 2 days in the dark at 4°C to break

dormancy (stratification) and transferred to 16/8 hr photoperiod at 120 mol m−2

s−1) at 21°C. After one week germination rate was evaluated. Four batches of

100 seeds were analyzed per line.

RNA extraction, cDNA library construction and sequencing

Total RNA was extracted with TRIZOL reagent (Invitrogen) according to the

manufacturer’s instructions, treated with DNAseI and purified with RNAeasy

Plant Mini Kit (Qiagen). A cDNA library was constructed using 1μg of total RNA

and the SMART™ cDNA Library Construction Kit (Clonthech) with oligo dT

priming. PCR-amplified cDNAs were ligated into pCR®II-TOPO (Invitrogen) and

introduced into E.coli. Insert amplification was performed by PCR using the

primers 5’-GGAAACAGCTATGACCATGATTACG-3’ and 5’–

GTCACGACGTTGTTAAACGACGGC–3’.



19

Single-pass sequencing was performed from the 5’ end of the selected PCR

products using the primer 5’–GTATCAACGCAGAGTCG–3’ at the Sequencing

Service (CRAG, Barcelona). The EST sequences (Accession numbers

AM937535 to AM939368) were compared against the public DNA and protein

databases using BLASTN, BLASTX and TBLASTX (Altschul et al., 1997) at

NCBI (http://www.ncbi.nlm.nih.gov/BLAST/) and CoGeBLAST

(http://genomevolution.org/CoGe/CoGeBlast.pl). The functional category

classification was derived from the BLAST results based in GO database

(http://geneontology.org/).

Construction, hybridization and data analysis of cDNA macroarrays

cDNA inserts were amplified by PCR using the primers: 5’-

ACTAGTTAATACGACTCACTATAGGGGGCCGAGGCGGCCGACATGTT-3’ and

5’-CCATGGTATTAGGTGACACTATAGAACGCAGAGTGGCCATTACGGCC-3’ in

96-well plates, their concentration adjusted to 100 ng µl-1 and denatured with

0.2 N NaOH. Spotting was done with a 96 pin Multi-Print Arrayer (V&P Scientfic)

using arrays of 36 x 24 spots and 200 nl hanging pins. Spots were printed onto

Hybond-N+ nylon membranes (Amersham). After air drying, membranes were

cross-linked by uv radiation at 120 μJ sec-1 for 1 min using Stratalinker

(Stratagene) and stored at 4˚C until usage. Hybridization probes were prepared

by the retrotranscription of 10 µg total RNA in presence of [α-32P]dCTP

annealed with 1 µg oligo(dT)12-18 (Invitrogen), using Supercript II RT

(Invitrogen) for 70 min at 42ºC. The RNA strand was hydrolyzed by adding 1 μl

5 M NaOH (37°C for 15 min) and the reaction was neutralized by adding 1 μl 5

M HCl and 5 μl 1 M Tris-HCl (pH 7.0). Probes were purified using Bio-Spin P-30

columns (Amersham). Pre-hybridization was done with MicroHyb hybridization

buffer (Invitrogen), 10% dextrane sulphate and 100 μg ml-1 denatured salmon

sperm DNA (42ºC for 2 hours). Hybridizations were done in the same buffer

(42ºC for 18 hours). Washing was done in 2 x SSC / 0.1% SDS twice for 30 min

at 65ºC. Washed membranes were exposed to a phosphoimager screen (GE

Heathcare) for 3 days and the screens scanned in a phosphoImager (Molecular

Imager, Bio-Rad). Hybridizations were done in triplicate. Image files were

imported into the Quantity-One program (Bio-Rad) for spot quantification.



20

Normalization was done based on the average intensities of seven

housekeeping genes: actin (AM937802), heat shock protein Hsp70

(AM939105), elongation factor 1-alpha (AM938565), translation initiation factor

5A (AM937959), polyubiquitin containing 7 ubiquitin monomer (AM938744),

beta-7 tubulin; (AM938025) and ubiquitin extension protein 2 (AM938244).

Statistical analysis was done using the t-student test (p < 0.05).

In silico analyses

Expression data on maize genes was obtained in Maize GDB

(http://www.maizegdb.org/expression/expr_tools/expr_tools.php) and maize

eFP Browser (http://bar.utoronto.ca/efp_maize/cgi-

bin/efpWeb.cgi?dataSource=Sekhon_et_al; Sekhon et al., 2011). Expression

data on Arabidopsis genes was obtained in the Arabidopsis eFP Browser

(http://bbc.botany.utoronto.ca/efp; Winter et al., 2007), Genevestigator

(https://www.genevestigator.ethz.ch/; Zimmermann et al., 2004) and

AtGeneExpress Visualization Tool (http://jsp.weigelworld.org/expviz/expviz.jsp;

Schmid et al., 2005). Protein Motif analyses were performed in SMART

(http://smart.embl-heidelberg.de/; Letunic et al., 2012), TargetP

(http://www.cbs.dtu.dk/services/TargetP/; Emanuelsson et al., 2000), ChloroP

(http://www.cbs.dtu.dk/services/ChloroP/; Emanuelsson et al., 1999), SignalP

(http://www.cbs.dtu.dk/services/SignalP/; Petersen et a., 2011) and TMpred

(prediction of transmembrane regions and orientation;

http://www.ch.embnet.org/software/TMPRED_form.html; Hofmann and Stoffel,

1993). Calculation of the hydropathy was done using ProtScale

(http://www.expasy.ch/tools/protscale.html; Gasteiger et al., 2005) based on the

Kyte & Doolittle index (Kyte and Doolittle, 1982).

The sequence of the maize OBAP1 protein was used as a query for the BLASP

and TBLASTN programs against several sequence databases including

GeneBank (http://blast.ncbi.nlm.nih.gov/), DOE Joint Genome Institute

(http://www.phytozome.net/search.php?show=blast), Plant GDB

(http://www.plantgdb.org/cgi-bin/blast/PlantGDBblast), GrainGenes

(http://wheat.pw.usda.gov/GG2/index.shtml), The Gene Index Project



21

(http://compbio.dfci.harvard.edu/tgi/), Rice Genome Annotation Project

(http://rice.plantbiology.msu.edu/), DFCI Maize Gene Index

(http://compbio.dfci.harvard.edu/tgi/cgi-bin/tgi/gimain.pl?gudb=maize), TIGR

Rice Genome Annotation Project (http://tigrblast.tigr.org/euk-

blast/index.cgi?project=osa1), The Maize Genome Browser

(http://www.maizesequence.org/Multi/blastview) and Arabidopsis Information

Resource (TAIR) (http://www.arabidopsis.org/). Taxonomical classification was

determined according to the NCBI Taxonomy Homepage

(http://www.ncbi.nlm.nih.gov/Taxonomy/taxonomyhome.html/). Amino acid

sequences were aligned using clustalw (Pole Bioinformatique Lyonnais:

http://npsa-pbil.ibcp.fr/cgi-

bin/npsa_automat.pl?page=/NPSA/npsa_clustalw.html). Phylogenetic tree was

constructed using Neighbor-Joining method using Treecon program (Van de

Peer and De Wachter, 1994).

Quantitative real-time RT-PCR

Total RNA was extracted from different tissues using the RNeasy plant mini kit

(Qiagen) according to the manufacturer’s instructions. From 2 µg of total RNA,

DNA was synthesized by the Omniscript RT kit (Qiagen) using an oligo-dT

primer according to the manufacturer’s instructions. cDNA samples were

normalized by OD260 and PCR reactions were set up in a LightCycler 480

system (Roche Diagnostics, Penzberg, Germany) using SYBR® Premix Ex

Taq™ (TaKaRa) using 200nM of the appropriate primers pairs and 2 µL of cDNA

template up to a volume of 20 μl. Reactions included an initial denaturation step

at 95ºC for 30 s, followed by 50 cycles of 95 ºC for 5 s, 60°C for 20 s and 72 °C

for 20 s. Melting curve analysis was performed on each sample to ensure single

amplicon specificity. Actin was used as reference endogenous control for

normalization purposes (primers 5’-TACCCAACTAAGCGCATGCC- 3’ , 5'-

GCATCTGAATCACGAAGCAGG-3'). Relative quantification was performed

using standard curves from serial dilutions of cDNA. The efficiency for all genes

tested were >99 %. All PCR reactions were performed in triplicate for each RNA

sample. Relative quantification analyses were performed using the LightCycler

480 Software. Gene-specific primers were designed using the Primer Express



22

software (Applied Biosystems, http://www.appliedbiosystems. com/). Specific

primers used were: for Obap1 5’-CCGGTGTTCCTGATCTTCA-3’, 5’-

GCACGTGAGCGAGGACAG-3’ and for oleosin1 5’-

TCCCGCAGATCATGATGGT-3’, 5’-CGAACTTCTTCTCCACACAGTCA-3’.

Construction of transient expression clones and sub-cellular localization

DNA sequences encoding the complete coding regions of maize OBAP1 and

Oleosin2 genes were amplified by PCR using the following primers: 5’-

CACCATGGCGTCGTCGTGCCAGAAC-3’ and 5’-

TTTAGTGAAGACCCTCCCGCC-3’, for OBAP1, and 5’-

CACCATGGCGGACCGTGACCGCAGC-3’ and 5’-

TTTCGAGGAAGCCCTGCCGCCGCCC-3’, for oleosin2. The resulting coding

fragments were cloned into the Gateway® binary vectors pYL-CFPct and pYL-

YFPct, respectively, to be driven by a 35S promoter of Cauliflower mosaic virus

(Rubio et al., 2005). Partial obap1 coding regions were obtained using the

following primers: 5’-CACCATGGCGTCGTCGTGC-3’ and

5’GTCCACCTGCCAGAAGTGGATGGT-3’, for N-OB, and 5’-

CACCATGGCGTCGTCGTGC-3’ and 5’- GTCCACCTGCCAGAAGTGGATGGT-

3’, or C-OB. The resulting fragments were cloned into the Gateway® binary

vector pYL-YFPct.

Plant transformation

The floral-dip method was used for Arabidopsis stable transformation (Clough

and Bent, 1998). For transient transformation, Nicotiana benthamiana leaves

were agro-infiltrated by syringe method into the abbatial side of 3 to 5-week-old

leaves (Voinnet et al., 2003). The binary plant vectors produced were mixed

together with the HC-Pro silencing suppressor construct (Goytia et al., 2006).

Three days after agro-infection, transformed leaves were observed.



23

Confocal laser-scanning microscopy

Confocal laser-scanning microscopy images were acquired using an Olympus

Fluoview FV10i inverted confocal laser-scanning microscope using water

immersion 60x objective. Fluorophore emissions were collected sequentially in

double-labeling experiments. Single-labeling experiments showed no detectable

crossover at the settings used for data collection. CFP was excited at 405 nm

and was detected with the 460–500 nm filter set. YFP was excited at 515 nm

and was detected with the 530–560 nm filter set. Nile Red was excited at 559

nm and was detected with the 570–670 nm filter set. For Arabidopsis embryo oil

body staining, Arabidopsis embryos were prepared from freshly harvested

seeds and oil bodies were stained with Nile Red as previously described

(Greenspan et al., 1985).

Immunodetection of OBAP1a

The peptide antibody against Arabidopsis OBAP1a was produced by Abyntek

Biopharma S.L. using the C-terminal peptide CEVDIKPVESVPRVFV. For

immunodetection of OBAP1a, Arabidopsis tissues were homogenized in protein

extraction buffer (50 mM Tris, pH 8.0, 2% SDS, and protease inhibitor cocktail).

Aliquots of the extracts were subjected to SDS-PAGE and transferred to

nitrocellulose membranes. Proteins were probed against the anti-OBAP1a

antibody and detected by alkaline phosphatase.

Electron microscopy

Isolated embryos were fixed for 2 h in 2% paraformaldehyde (w/v) and 2.5%

glutaraldehyde (v/v) in phosphate buffer 0.1M, pH 7.4. Following fixation, the

embryos were washed with phosphate buffer 0.1M, pH 7.4 for 10 min and

resuspended in 1% paraformaldehyde (w/v) for 2 h and rinsed three times with

phosphate buffer. Embryos were post-fixed for 2 h with 1% osmium tetroxide

(w/v) containing 0.8% potassium hexocyano ferrate (w/v) prepared in phosphate

buffer, followed by four washes with deionized water and sequential dehydration

in acetone. All procedures were performed at 4ºC. Samples were embedded in



24

Spurr´s resin and polymerized at 60ºC for 48 h. Ultrathin sections (70 nm) were

obtained with a diamond knife (45º, Diatome, Biel, Switzerland). Sections were

stained with uranyl acetate and Reynolds lead citrate for 10 min and examined

with a Jeol 1400 transmission electron microscope (Jeol Ltd, Tokyo, Japan)

equipped with a Gatan Ultrascan ES1000 CCD Camera. Embryos from three

individuals were used in each analysis.

For gold immunolabeling ultrathin sections were incubated in phosphate buffer

with anti-OBAP1a antibody at 4ºC overnight in a humidified chamber. The grids

were washed with 0.5 % Tween-20 (v/v) in PBS and incubated in the blocking

solution with a secondary anti-rabbit antibody coupled to 10 nm gold particles

for 1 h at room temperature. Grids were washed five times with 0.5% Tween-20

(v/v) in PBS 10 min, washed with PBS and finally with deionized water. Grids

were stained with uranyl acetate and Reynolds lead citrate solutions.

Subcellular fractioning

5 g of dry seeds was homogenized in buffer (0.4M sucrose, 10 mM KCl, 1 mM

MgCl2, 1 mM EDTA, 100 mM Hepes pH 7.5, 0.1 mM PMSF) with a mortar and

pestle at 4ºC. Homogenates were filtered through Miracloth and centrifuged at

13.000 rpm for 2 min to remove debris. Crude homogenates were then

centrifuged at 13.000 rpm for 10 min at 4ºC. The floating oil body fraction was

removed and resuspended in 200 µl 50 mM Tris-HCl (pH 7.2) buffer containing

8 M urea and the non-floating fraction was stored as cytosolic fraction. The

resuspended floating oil body fraction was diluted with 1 ml of buffer and oil

bodies were recovered by centrifugation as before. This step was repeated

twice. Both cytosolic and oil body fractions were delipidated with 5 vol of

acetone. Proteins were recovered by centrifugation at 13.000 rpm for 2 min. Dry

pellets were resuspended in 200µl 50 mM Tris-HCl pH 7.2 and 0.1% SDS.

Protein concentration was determined using protein assay from BioRad

(Hercules, CA). 10 µg of protein was loaded per lane on 12% SDS-PAGE gel.



25

TAG analysis

Lipids were extracted from 50 Arabidopsis seeds as previously described by

Burgal et al. (2008), not including 1,1,1-13C-triolein as an internal standard. TAG

purification was carried out by one-dimensional thin-layer chromatography

according to Hernández et al. (2008). Fatty acid methyl esters from TAGs were

produced by acid-catalyzed transmethylation (Garcés and Mancha, 1993) and

analyzed by gas chromatography using a 7890A (Agilent technologies, Santa

Clara, CA, USA) fitted with a capillary column (30 m length; 0.25 mm i.d.; 0.20

µm film thickness) of fused silica (Supelco, Bellafonte, PA, USA) and a flame

ionization detector. Hydrogen was used as carrier gas with a linear flux of 1.34

ml min-1 and a split ratio of 1/50. The injector and detector temperature was

220 ºC, and the oven temperature 170 ºC. Heptadecanoic acid was used as

internal standard to calculate the lipid content of the samples.

SUPPLEMENTAL MATERIAL

Table S1.- EST sequences from immature scutellum library

Table S2.- Maize genes represented in the EST sequences from immature

scutellum library

Table S3.- Functional categories of the maize genes represented in the EST

sequences from immature scutellum library

Table S4.- Array hybridization results.

Table S5.- Species whose genomes contain genes coding for OBAP proteins.

Figure S1.- Polypeptide sequence alignment of OBAP proteins. Sequence were

aligned using the ClustalW algorithm.

Figure S2.- Neighbor-joining tree of OBAP proteins from different species.



26

Figure S3.- Expression analysis of Arabidopsis obap1a gene (At1g05510)

based on the eFP Browser database information. Relative expression levels of

Arabidopsis obap1a gene in different organs and developmental stages.

Figure S4.- Mechanism of oil body association. A-C, Kyte-Doolittle hydropathy

index plots for different maize oil body associated proteins calculated with

window size of 15 amino acids. Each signal in the X-axis represents 10 amino

acids. A, oleosin1 (NP_001105338.1); B, caleosin (NP_001151906); C,

steroleosin (NP_001152614.1).

ACKNOWLEDGMENTS

We thank Dr. Juanjo Lopez-Moya for providing p35S:HcPro, Dr. Vicente Rubio

and Dr. Salomé Prat for provide us with the pVR-eCFPCt and pVR-YFPCt

vectors. We thank Maria Coca for provide us with the oleosin2-CFP construct.

We thank Montse Amenós (Core Microscopy Facility, CRAG) and Alejandro

Sánchez (Electronic Microscopy Facility, UAB) for technical assistance.

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FIGURE LEGENDS

Figure 1.- Unrooted neighbor-joining tree based on aligned protein sequences

of OBAP proteins from different species.

Figure 2. Relative abundance of maize obap1 and oleosin1 genes normalized to

actin in different organs and developmental stages measured by quantitative

RT-PCR. The standard deviation of three repeats is shown. A, C and E, obap1

mRNA accumulation. B, D and F, oleosin1 mRNA accumulation. A and B, total



34

RNA from dissected parts of maize kernels 30 days after pollination (Scut,

scutellum; Axis, embryo axis; Endos, endosperm), and dissected parts of 15

days old seedlings (leaf and root). C and D, total RNAs extracted from immature

embryos at different days after pollination (dap). E and F, total RNAs extracted

from dissected scutellum of embryos at 30 days after pollination (30 dap), from

dry seeds (ds), and at different days after imbibition (dai).

Figure 3.- Arabidopsis OBAP1a protein. A, Immunoblot of homogenates from

Arabidopsis seeds at different stages of development (S1, globular stage, to S6,

dry seeds), and from Arabidopsis mature leaves (Le), and stems (St). B,

Immunoblot of homogenates from Arabidopsis seeds and seedlings at different

days after imbibition.

Figure 4.- Subcellular localization of maize OBAP1 in tobacco cells. Confocal

projection images of transiently transformed Nicotiana benthamiana leaf

epidermal cells. A, tobacco cell transformed with OBAP1-YFP. B, tobacco cell

transformed with YFP. C, non transformed tobacco cell stained with Nile Red.

D, tobacco cell transformed with OBAP1-YFP. E, the same transformed tobacco

cell stained with Nile Red. F, co-localization of OBAP1-YFP and Nile Red. G,

tobacco cell transformed with Oleosin2-CFP. H, the same tobacco cell

transformed with OBAP1-YFP. I, co-localization of Oleosin2-CFP and OBAP1-

YFP; arrows indicate ER-like structures; triangles indicate OB-like structures. In

A-F the bar corresponds to 10 μm. In G-I the bar corresponds to 2 μm.

Figure 5.- Ultraestructural localization of OBAP1 in rapeseed embryos.

Immunolabelling of oil bodies with anti-OBAP1 serum. The position of oil bodies

is indicated (ob). The black or white arrows indicate the position of the gold

particles. A, B, C and D images correspond to different rapeseed embryo

samples treated similarly.

Figure 6.- Western–blot analysis of the total protein (TOT), oil body fraction

(OB) and soluble protein fraction (SOL) of Arabidopsis mature seed extract

using an anti-AtOBAP1a antibody. The arrow indicates the expected position of

AtOBAP1a.



35

Figure 7.- Mechanism of oil body association of OBAP1. A, Kyte-Doolittle

hydropathy index plot for maize OBAP1 protein calculated with window size of

15 amino acids. Each signal in the X-axis represents 10 amino acids. B,

Schematic representation of the two fragments of OBAP1 protein fused to YFP.

C-E, Confocal projection images of transiently transformed Nicotiana

benthamiana leaf epidermal cells. C, Tobacco cell transformed with YFP. D,

Tobacco cell transformed with N-OBAP1-YFP (N-OB). E, Tobacco cell

transformed with C-OBAP1-YFP (C-OB). The bar corresponds to 8 μm.

Figure 8.- Arabidopsis AtOBAP1a mutants and interference lines. A, Schematic

representation of the position of the T-DNA insertions in the mutant lines 1A1

and 1A2 respect to the Arabidopsis obap1a gene. The arrow indicates the ATG

start codon. B, Immunoblot of homogenates from mature seeds of Arabidopsis

wild type (wt, Col-0), insertion mutants 1A1 and 1A2, and from the RNA

interference line Ag1, with anti-OBAP1a antibody. C, Germination rates of the

seeds of the different lines. Standard deviations are indicated. D,

Tryacylglygerol content of seeds of the different lines. E, Fatty acid composition

of TAGs of the different seed lines. Standard deviations are indicated.

Figure 9.- OBAP1 deficiency causes enlargement of oil bodies in embryos. (A-

C) Confocal projection images of Arabidopsis mature embryos stained with Nile

Red: A, Col-0; B, T-DNA insertion mutant 1A1; C, RNA interference line Ag1. (D-

I) Electron micrographs of Arabidopsis mature seeds: D-E, Col-0; F-G, T-DNA

insertion mutant 1A1; H-I, RNA interference line Ag1. Some of the oil bodies are

indicated by red arrows. In D-I the bar corresponds to 2 μm except in H that

correspond to 1 μm.



Monocots 1

Dicots 1

Conifers 1

Primitive plants 1Monocots 2

Dicots 2

Conifers 2

Primitive plants 2

BacteriaFungi

Figure 1.- Unrooted neighbor-joining tree based on aligned protein sequences of OBAP proteins from different species.



Figure 2. Relative abundance of maize obap1 and oleosin1 genes normalized to actin in different organs and developmental stages measured by quantitative RT-PCR. The standard deviation of three repeats is shown. A, C and E, obap1 mRNA accumulation. B, D and F, oleosin1 mRNA accumulation. A and B, total RNA from dissected parts of maize kernels 30 days after pollination (Scut, scutellum; Axis, embryo axis; Endos, endosperm), and dissected parts of 15 days old seedlings (leaf and root). C and D, total RNAs extracted from immature embryos at different days after pollination (dap). E and F, total RNAs extracted from dissected scutellum of embryos at 30 days after pollination (30 dap), from dry seeds (ds), and at different days after imbibition (dai).



2520

S1 S2 S3 S4 S5 S6 Le St

2520

0 1 2 3 4 5 6 7

A

B

Figure 3.- Arabidopsis OBAP1a protein. A, Immunoblot of homogenates

from Arabidopsis seeds at different stages of development (S1, globular

stage, to S6, dry seeds), and from Arabidopsis mature leaves (Le), and

stems (St). B, Immunoblot of homogenates from Arabidopsis seeds and

seedlings at different days after imbibition.



A

D

B

E F

Figure 4.- Subcellular localization of maize OBAP1 in tobacco cells. Confocal projection images of transiently transformed Nicotiana benthamiana leaf epidermal cells. A, tobacco cell transformed with OBAP1-YFP. B, tobacco cell transformed with YFP. C, non transformed tobacco cell stained with Nile Red. D, tobacco cell transformed with OBAP1-YFP. E, the same transformed tobacco cell stained with Nile Red. F, co-localization of OBAP1-YFP and Nile Red. G, tobacco cell transformed with Oleosin2-CFP. H, the same tobacco cell transformed with OBAP1-YFP. I, co-localization of Oleosin2-CFP and OBAP1-YFP; arrows indicate ER-like structures; triangles indicate OB-like structures. In A-F the bar corresponds to 10 μm. In G-I the bar corresponds to 2 μm.

C

HG I



A B

C D

200 nm 100 nm

100 nm100 nm

Figure 5.- Ultraestructural localization of OBAP1 in rapeseed embryos. Immunolabelling of oil

bodies with anti-OBAP1 serum. The position of oil bodies is indicated (ob). The black or white

arrows indicate the position of the gold particles. A, B, C and D images correspond to different

rapeseed embryo samples treated similarly.

obob

ob

ob

ob

obob

ob

ob

ob

obob ob

ob

ob

ob

ob

ob

ob

ob

ob

ob

ob

ob

ob

obob

ob

ob

ob



TOT OB SOL

Figure 6.- Western–blot analysis of the total protein (TOT), oil body fraction (OB) and soluble protein fraction (SOL) of Arabidopsis mature seed extract using an anti-AtOBAP1a antibody. The arrow indicates the expected position of AtOBAP1a.



N-OB C-OB

N-OB C-OB

+3

+2

+1

0

-1

-2

-3

A

B

C D

Figure 7.- Mechanism of oil body association of OBAP1. A, Kyte-

Doolittle hydropathy index plot for maize OBAP1 protein

calculated with window size of 15 amino acids. Each signal in the

X-axis represents 10 amino acids. B, Schematic representation

of the two fragments of OBAP1 protein fused to YFP. C-E,

Confocal projection images of transiently transformed Nicotiana

benthamiana leaf epidermal cells. C, Tobacco cell transformed

with YFP. D, Tobacco cell transformed with N-OBAP1-YFP (N-

OB). E, Tobacco cell transformed with C-OBAP1-YFP (C-OB).

The bar corresponds to 8 μm.

E



1A11A2A

B 1A1 1A2 wt Ag12520

C 100

0

25

50

75

% g

erm

inat

ion

1A2 1A1 Ag1wt

Figure 8.- Arabidopsis AtOBAP1a mutants and interference

lines. A, Schematic representation of the position of the T-DNA

insertions in the mutant lines 1A1 and 1A2 respect to the

Arabidopsis obap1a gene. The arrow indicates the ATG start

codon. B, Immunoblot of homogenates from mature seeds of

Arabidopsis wild type (wt, Col-0), insertion mutants 1A1 and 1A2,

and from the RNA interference line Ag1, with anti-OBAP1a

antibody. C, Germination rates of the seeds of the different lines.

Standard deviations are indicated. D, Tryacylglygerol content of

seeds of the different lines. E, Fatty acid composition of TAGs of

the different seed lines. Standard deviations are indicated.

D 5

0

1

2

3

1A1 1A2 Ag1wt

µg

TAG

/see

d 4

E

16:0 16:1 18:0 18:1 18:2 18:3 20:0 20:1 20:2 22:1

wt 1A1 1A2 Ag1

Fatty acid species



A B C

D

F

H

E

G

I

Figure 9.- OBAP1 deficiency causes enlargement of oil bodies in embryos. (A-C)

Confocal projection images of Arabidopsis mature embryos stained with Nile Red:

A, Col-0; B, T-DNA insertion mutant 1A1; C, RNA interference line Ag1. (D-I)

Electron micrographs of Arabidopsis mature seeds: D-E, Col-0; F-G, T-DNA

insertion mutant 1A1; H-I, RNA interference line Ag1. Some of the oil bodies are

indicated by red arrows. In D-I the bar corresponds to 2 μm except in H that

correspond to 1 μm.



obap, a new family of oil body-associated proteins author to whom

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